Physiological responses to a changing winter climate in an early spring‐breeding amphibian

Abstract Climate change is swiftly altering environmental winter conditions, leading to significant ecological impacts such as phenological shifts in many species. As a result, animals might face physiological mismatches due to longer or earlier activity periods and are at risk of being exposed to late spring freezes. Our study points for the first time to the complex physiological challenges that amphibians face as a result of changing thermal conditions due to winter climate change. We investigated the physiological responses to a period of warmer winter days and sudden spring freeze in the common toad (Bufo bufo) by acclimating them to 4°C or 8°C for 48 h or exposing them to 4°C or −2°C for 6 h, respectively. We assessed the daily energy demands, determined body condition and cold tolerance, explored the molecular responses to freezing through hepatic tissue transcriptome analysis, and measured blood glucose levels. Toads acclimated to higher temperatures showed a higher daily energy expenditure and a reduced cold tolerance suggesting faster depletion of energy stores and the loss of winter acclimation during warmer winters. Blood sugar levels were higher in frozen toads indicating the mobilization of cryoprotective glucose with freezing which was further supported by changed patterns in proteins related to glucose metabolism. Overall, our results emphasize that increased thermal variability incurs physiological costs that may reduce energy reserves and thus affect amphibian health and survival. This might pose a serious threat to breeding adults and may have subsequent effects at the population level.


| INTRODUC TI ON
Understanding the responses and adaptability of wildlife to climate change is still a major challenge in ecological research.To date, most of the research exploring the impacts of climate change on animals has focused on effects of extreme heat events as well as on how animals might cope with environmental temperatures close to their upper thermal limits (e.g., Comte & Olden, 2017;Morley et al., 2019;Sunday et al., 2019).However, in many northern temperate ecosystems, shifts in temperature and precipitation patterns have been, and are expected to be, more pronounced during winter compared to summer (Lee et al., 2023;Williams et al., 2015).Winter climate change is driving complex changes in both thermal means and temperature extremes with a generally increasing trend (rev. in Marshall et al., 2020).Particularly at high latitudes, climate is warming significantly faster in winter than in summer (Cohen et al., 2014;Wallace et al., 2014).Furthermore, some regions are experiencing unusual cold spells or extreme weather events as a result of climate-induced changing circulation patterns, making winters more variable in addition to warmer (Stendel et al., 2021).
A common ecological consequence of winter climate change across diverse taxa is phenological shifts in critical life-history events (Cohen et al., 2018;Parmesan & Yohe, 2003;Visser & Both, 2005;Walther et al., 2002).These include an accelerated onset of the growing season for many plants (Cleland et al., 2007), an advanced return of in migratory birds (Gordo, 2007), the anticipated emergence of many dormant insect or mammal species (Forrest, 2016;Wells et al., 2022), as well as an earlier initiation of the breeding season (Green, 2017).Among all taxa, amphibians have been shown to reveal the strongest average shift towards earlier breeding (Parmesan, 2007), with >60% of all populations studied doing so (Li et al., 2013).However, the magnitude of phenological response is known to be species-and population-specific, and to vary with latitude (Parmesan, 2007;While & Uller, 2014).
With a shifted breeding phenology induced by winter warming, adult amphibians are likely experiencing an extended or earlier annual activity period.Since temperate amphibians rely on internal energy reserves (i.e., the fat body) during overwintering, warmer winter temperatures as well as extended or earlier activity periods might increase metabolic costs and drain such energy reserves (Arambourou & Stoks, 2015;Reading, 2007) with consequences for body condition and survival (Jørgensen, 1986;Reading & Clarke, 1995).In addition to warming, climate change will also increase the occurrence of extreme weather events such as sudden frost in spring known as false springs (Ault et al., 2013;Meehl et al., 2000), thereby exposing amphibians to the increased risk of cold stress.Despite an increased risk of being frozen to death or frost damage, experiencing cold stress after the reproductive season has begun is physiologically costly (Ruthsatz et al., 2022).This cold stress has many adverse effects including reduced energy reserves, comprised immune system functioning, and decreased fecundity (rev. in Liu et al., 2018), and might therefore pose a serious threat to breeding amphibians.
Amphibians have developed several responses to cope with cold stress such as the selection of microhabitats with more favorable temperatures to mitigate cold exposure (Boutilier et al., 2000;Sinsch & Leskovar, 2011) or physiological adaptations to endure cold conditions (Costanzo & Lee Jr, 2013;Storey & Storey, 1988).If behavioral thermoregulation is constrained, such as through the occurrence of sudden frost, freeze protection or tolerance through the accumulation of cryoprotectants, such as plasma glucose derived from liver glycogen catabolism, glycerol, or urea, might contribute to withstanding extreme cold exposure (do Amaral et al., 2018;Storey & Storey, 1985, 2017).However, if energy reserves are drained due to warmer winter temperatures as well as extended or earlier activity periods, the ability to successfully implement an effective response to cold stress might be impaired (Marshall et al., 2020).
Consequently, increases in mean temperature could potentially increase the susceptibility to extreme weather events such as sudden frost.Nonetheless, the physiological challenges and potential impacts of changing winter climate on amphibians are still not well understood.
In this study, we investigated the physiological responses of early spring-breeding temperate amphibians to changing thermal conditions resulting from winter climate change, using the common toad (Bufo bufo) as a model species.Specifically, we aimed to determine metabolic costs resulting from higher winter temperatures combined with exploration of cryoprotective mechanisms in response to sudden (night) freeze.To simulate warmer winter days and spring freeze, toads were acclimated to 4°C or 8°C for 48 h (i.e., experiment 1) or exposed to either 4°C or −2°C for 6 h (i.e., experiment 2), respectively.We measured the standard metabolic rate to assess daily energy demands and determined body condition, cold tolerance as well as glucocorticoid hormone levels as a proxy for physiological stress in toads acclimated to different winter temperatures.To explore the molecular responses to freezing, we used a transcriptomelevel investigation of hepatic tissue, and measured blood glucose level.We predicted that toads acclimated to warmer winter temperature would experience reduced cold tolerance, higher levels of physiological stress, and increased metabolic rate, resulting in higher mass loss and depletion of energy budgets compared to the control group.We further predicted that toads facing false springs would respond with the mobilization of cryoprotective glucose and reveal differentially regulated hepatic pathways related to cryoprotectant mobilization and stress tolerance.

| MATERIAL S AND ME THODS
We ran two experiments to investigate the physiological responses to warmer winter days (i.e., 2.2 Experiment 1: Warmer winter scenario) and sudden spring freeze (i.e., 2.3 Experiment 2: Spring freeze scenario) in the widespread anuran B. bufo.This early spring-breeding amphibian was selected as a model species in this study because its spring migratory and spawning behavior directly exposes it to changes in temperature caused by winter climate change, and its reproductive success can be strongly influenced by these changes (Reading, 1998).

| Field work and animal collection
The fieldwork was carried out at the locality Kleiwiesen (52.328 N,10.582E) near Braunschweig, Germany, after the migration of B. bufo had started on March 17th and 18th, 2022 (Table S1).This site sustains a large population of B. bufo, which breed in a shallow part of one pond, partly covered with dense reeds.In total, we collected 40 toads (i.e., n experiment1 = 30; n experiment2 = 10) by hand wearing nitrile gloves and kept them individually in small plastic boxes containing wet tissue to avoid dehydration (Tables S1 and S2).Special care was taken to capture only male toads on their way to the breeding pond to avoid any impact on the population, considering that many female toads typically have just one opportunity to spawn in their lifetime (Kuhn, 1994).Mean ambient temperature during the toad collection was 3.72°C (Table S1).Therefore, we considered 4°C as temperature threshold for the start of the migratory activity in male toads (Gittins et al., 1980;but see: Sinsch, 1988) and used this temperature as control temperature in both experiments.After collection, the toads were transported to the laboratory on the day of collection, where their snout-vent-length (in cm; SVL) and body mass (in g) were measured.SVL was measured with a caliper (to the closest 0.05 cm) and body mass was determined with an electronic balance (to the nearest 0.01 g; Sartorius A200 S; Table S1).As digestion is accompanied by an increase in metabolic rate, animals were fasted for 48 h (i.e., during the thermal acclimation period) before the physiological measurements (Orlofske & Hopkins, 2009).This fasting mimics natural conditions as during the migration to the breeding ponds, spring breeding amphibians rely on their internal fat storages (i.e., fat bodies; 22).

| Animal husbandry and acclimation conditions
To test how a period of warmer winter days affects the physiology of B. bufo, 30 animals were randomly split into two groups and acclimated to 4°C and 8°C for 48 h, respectively, using two different refrigerators.Acclimation temperatures were chosen in order to simulate thermal conditions during early spring (i.e., 4°C) and warmer conditions (i.e., 8°C) as predicted under changing global climate following the approach of Podhajský & Gvoždík (2016).Refrigerators were set to the respective acclimation temperature and equipped with additional digital thermometers (TFA LT-102, Conrad electronics; precision: ±0.5°C).Animals were housed individually in plastic boxes (10 cm × 20 cm × 15 cm) with moist tissue to prevent dehydration.Tissues were changed daily and new water was added to keep the boxes clean.

| Physiological measurements
After the 48-h acclimation period, the following physiological parameters were determined: body condition, water-borne corticosterone (CORT) release rates, standard metabolic rate (SMR), daily energy expenditure (DEE), and cold tolerance (Table S1).After the physiological measurements, all 30 toads were kept at 4°C for an additional 48 h and fed medium-sized house crickets (Acheta domesticus) ad libitum to allow them to recover and feed before we returned them to the collection site.

Body condition
Body condition was determined directly after collection as well as after 48 h (i.e., acclimation duration) to examine if the body condition changed during this period.The scaled mass index (SMI) was used to determine body condition from body mass and SVL following (MacCracken & Stebbings, 2012;Peig & Green, 2009).The SMI slope is calculated from the regression of log transformed SVL and log transformed mass.

CORT assays
After 48 h of acclimation to either 4°C or 8°C, we measured CORT levels in 30 toads using the established water-borne assay protocol by Gabor et al. (2013).Briefly, toads were kept in freshly cleaned (EtOH) plastic containers with a volume of 480 mL containing 100 mL of aged and filtered tap water for 1 h.For each sampling batch (i.e., measurement of five toads at the same time), a control sample was run with the sampling water to account for any potential background hormonal traces (Ruthsatz et al., 2023).After taking the control samples the containers were placed in a water bath (Lauda ECO RE 2025 G; LAUDA DR.R. WOBSER GMBH & CO.KG) set to respective acclimation temperature (i.e., 4°C and 8°C).During the collection period, nobody was allowed to enter the room to avoid any disturbance of the animals (minimal noise and visual disturbances).After the hour-long sample collection period, we determined the body mass of each toad, which was followed by metabolism measurements.
All water hormone samples were stored at −25°C.We processed them in a random order within 1 month.Control samples (N = 6) were pooled before processing.Thawed samples were first filtered with Q8 Whatman filter paper to remove suspended particles, and then filtered through C18 solid-phase extraction columns (Oasis Vac Cartridge HLB 3 cc/60 mg, 30 μm; Waters Inc.) with a vacuum manifold (Visiprep Vacuum Manifold; Sigma-Aldrich).The manifold was cleaned before each use with 4 mL of HPLC-grade ethanol and 4 mL of nanopure water.The columns were returned to the −25°C freezer until hormones were eluted with 4 mL of HPLC-grade methanol with a vacuum manifold (Visiprep Vacuum Manifold; Sigma-Aldrich), which was cleaned with ethanol and nanopure water between batches again.During this process, samples were transferred into 5 mL Eppendorf tubes.Afterwards the methanol was evaporated using The hormonal levels were measured using DetectX Corticosterone ELISA (Enzyme Immunoassay) kits purchased from Arbor Assays (K014-H5; assay has a range of 19.53-5000 pg Corticosterone/mL).Control samples were run on each plate in duplicate.This assay has been previously validated for anurans such as R. temporaria (Burraco et al., 2017;Ruthsatz et al., 2023).Samples and kit reagents were brought to room temperature and vortexed before plating.We measured corticosterone concentration in duplicates for all samples on 96-well plates according to the kit's instructions.The plates were read with a Tecan Spark® Microplate Reader at 450 nm (Tecan).In total, we ran one plate.
Control samples and negative controls did not show CORT levels at detectable ranges.We used MyAssays online tools to calculate the hormonal concentration of samples (https:// www.myass ays. com/ arbor -assay s-corti coste rone-enzym e-immun oassa y-kit-impro ved-sensi tivity.assay ).Two positive control samples were run in duplicate on the plate to assess intra-assay variation.Intraplate variation was overall 1.74% (high: 2.68, low: 0.99).The coefficient of variation of duplicates for all samples was 2.72%.
Following Gabor et al. (2013), we multiplied CORT release rates (pg × mL −1 ) by the volume of the re-suspension solution (0.5 mL) and standardized values by dividing by the body mass of each individual, resulting in CORT release rates units being pg × g −1 × h −1 .

Metabolism and energy expenditure
Oxygen consumption was measured by closed respirometry.
Individuals were measured in airtight plastic containers with a volume of 480 mL at their respective acclimation temperature (i.e., 4°C and 8°C).In each container, a planar oxygen sensor spot (SP-PSt3-NAU-D5-YOP, PreSens Precision Sensing GmbH) was integrated, which was connected to a multichannel oxygen measuring system (Oxy-4 SMA; PreSens Precision Sensing GmbH) via a fiber optic sensor (Polymer Optical Fiber POF, PreSens Precision Sensing GmbH).
Prior to each trial, the O 2 fiber optic sensors were calibrated using air-saturated water and a factory-set zero oxygen calibration point

Cold tolerance
The cold tolerance was determined for each toad by using the loss of righting response as a proxy for the lower critical thermal tolerance (CT min ) according to the procedure of McCann et al. (2018).
The initial body temperature of the toads was measured by inserting a temperature probe (ThermoPro TP622 IP65 waterproof digital thermometer, ThermoPro DE; precision: ±0.3°C) into the animal's cloaca (i.e., T cloaca ).The toad was then placed inside a closed plastic container.The container was placed inside an insulated box that was filled with ice.During the cooling, the toad was gently removed from the box every 10 min and placed on its back on a flat surface.If the toad righted itself within 30 s, it was returned to the ice-filled box for further 10 min.After every 10 min the procedures of the measurements were repeated until the toad was no longer able to turn over within 30 s.The T cloaca of the toad at this point was recorded as its CT min .This measurement represents a biologically valid and ethically acceptable measure of an animal's ability to function (McCann et al., 2018;Taylor et al., 2021).After these procedures, the toads were placed back into their boxes and warmed to their initial acclimation temperature with a heating rate of 0.1°C per minute.All toads recovered quickly.

| Statistics
For all statistical tests Cran R (Version 4.1.1,R Development Core Team, 2022) for Windows was used.All plots were constructed using ggplot2 (Wickham, 2009).Statistical significance was accepted at α < .05.
All dependent variables in the models were log 10 -transformed and tested for autocorrelations using Spearman's rank correlation before the analyses (cor.testfunction in the corTest package; Yu et al., 2019).Consequentially, variables were included in statistical analyses when the correlation was significant but below the standard threshold of 0.7 or particularly essential for comparison with previous studies (i.e., SMR/CORT; Chin, 1998; Table S3).A1).Toads were handled wearing insulating gloves.
Immediately after freeze exposure, animals from both the freeze group and the control were euthanized by a sharp blow on the head, followed by rapid decapitalization following the protocol of the German animal protection law in an approved laboratory space.
Right after euthanizing, the liver was dissected free from surrounding tissue, aliquoted into triplicates, transferred to 1.5 mL Eppendorf tubes filled with RNAlater, snap frozen in liquid nitrogen, and stored at −80°C until transcriptomic analysis.

| Blood glucose level
In freshly euthanized toads, blood glucose levels (mg × dL −1 ) were determined in three toads per treatment using a portable kit (CONTOUR®NEXT, Ascensia Diabetes Care Deutschland GmbH) from the carotid artery.A Mann-Whitney U-test was used to compare blood glucose levels between toads exposed to freeze and the control group.

| Fat body analysis
Fat bodies of four toads were dissected, dry blotted, and weighed to the nearest 0.001 g, followed by drying at 50°C for 24 h.Gross energy content of dissected fat bodies in J was identified by bomb calorimetry (PARR 6100 bomb calorimeter, PARR Instruments Deutschland GmbH) with benzoic acid as calibration standard at the University of Hamburg.We lost one sample due to a misfire of the calorimeter.All results were converted from dry matter to fresh weight by correcting the weight loss of the fat body during drying.

| Transcriptomics
Total RNA was extracted with a phenol-chloroform precipitation protocol and measured with a NanoDrop 1000 (Peqlab Biotechnologie GmbH).The RNA samples were then prepared with the Illumina TruSeq stranded mRNA protocol.Afterwards, the samples were quantified with the Quant-iT™ dsDNA BR Assay Kit on a NanoDrop 3300 fluorometer.Equimolar amounts of the samples were pooled.
The complete library was run on an Agilent Bioanalyzer prior to being prepared for a NextSeq run, as recommended by Illumina.The library was sequenced on a NextSeq500 using the NextSeq 500/550 HighOutput Kit v2.5 2 × 75 bp cycles sequencing chemistry.
Subsequently, the quality and integrity of the assemblies were assessed through the Trinity tool Trinity.stats.For the differential expression (DE) analysis, transcript expression levels were quantified using the tool Salmon (Patro et al., 2017), which was integrated into the Trinity pipeline through the execution of the Perl script align_ and_estimate_abundance.pl.Following this, transcript count matrices were generated, and cross-sample normalization was performed using the abundance_estimates_to_matrix.plscript.
Differential expression analysis, conducted at both the gene and isoform levels, was carried out using DESeq2 (Love et al., 2014), also integrated into the Trinity pipeline.The p-value threshold was set to p < .01,and a minimum count threshold of two was applied.The DESeq output was then employed to create heatmaps for differentially expressed genes and isoforms using the R package pheatmap (Kolde, 2019).Additionally, volcano plots were generated by plotting the log 2 fold change (logFC) against the −log 10 (adjusted p-value) using the R package ggplot2.

Transcriptome annotation
The de novo assembly was annotated following the Trinotate pipeline (Bryant et al., 2017) by extracting predicted coding regions using TransDecoder (http:// trans decod er.github.io) and searching both the entire transcripts and the predicted coding regions separately by BLAST+ (Camacho et al., 2009) against the Uniprot Swiss-Prot database (UniProt Consortium, 2019), with a stringent expectation value (e-value) of 1 × 10 −5 , retaining only the best BLAST hits.The results were stored in an SQL database, with two distinct tables created for differentially expressed genes and isoforms.

Functional classification and gene ontology enrichment analysis
For functional classification, each BLAST hit was examined, and genes and isoforms with no or contradictory BLAST hits were filtered out.Subsequently, protein functions were identified by referencing the online UniProt database (https:// www.unipr ot.org/ ) and conducting a systematic literature review for potential connections between the identified proteins and their roles in freeze response. The

| Experiment 1: Warmer winter scenario
There was a significant difference in all physiological parameters between the toads in the 4°C and 8°C acclimation temperature treatment (Table S1 and Figures 1-3).Toads acclimated to 8°C revealed a 3.7-times higher CORT release rate (U = 0.00; p < .001) as well as a 27.1% higher SMR (U = 0.0; p < .001)and used 92.96J more energy per day (U = 45.00;p = .004).Mean (±SD) fat body mass and energy content were 0.243 g (±0.08) and 8652.68J, respectively.Based on the DEE of the toads at the different acclimation temperatures, mean fat body energy content would be sufficient to cover the energetic maintenance costs 5.26 day longer in toads kept at lower temperatures (i.e., 25.53 day at 4°C; 20.27 day at 8°C; Figure 1).
There was a significant difference in body mass and body condition before and after the 48-h-long acclimation period (Table S1 and of the toads lost body mass and body condition, whereas 26.61% revealed a higher body mass and body condition. Cold tolerance was significantly lower (i.e., CT min was higher) in toads acclimated to 8°C (U = 2.50; p < .001;N = 30; Figure 3).Mean (±SD) CT min was −0.167 (±0.13) and 0.34 (±0.14) in toads at 4°C and 8°C, respectively.

| De novo assembly and annotation
The de novo assembly was conducted using RNA-seq data obtained from both control (n = 5) and frozen toad (n = 5) samples.cDNA libraries were constructed from the liver tissue of B. bufo, resulting in a comprehensive assembly comprising 231,696 transcripts (see Table S4 for full assembly statistics).Employing DESeq2 for differential gene expression analysis revealed significant variations in both gene and isoform expression patterns between the freeze and control group (Figure 4 and Figures A2-A5).A total of 54 differentially expressed transcripts were identified at the gene level, while at the isoform level, 277 transcripts showed distinct expression profiles (Figure 4 and Figure A4).Subsequent annotation using Trinotate, coupled with a stringent filtering process for BLAST results (excluding transcripts F I G U R E 1 Gardner-Altman estimation plots displaying the mean difference between acclimation temperatures on (a) corticosterone (CORT) release rate (pg × g −1 × h −1 ), (b) standard metabolic rate (SMR, mL O 2 × h −1 × g −1 ), and (c) daily energy expenditure (J × day −1 ).Boxes and whiskers show 25th to 75th and 10th to 90th percentiles, respectively; black lines indicate the median.Dot = single data points.Blue boxes = toads acclimated to 4°C.Orange box = toads acclimated to 8°C.

(a) (b) (c)
lacking BLAST hits and those with contradictory outcomes), allowed us to assign names and functions to six proteins at the gene level and 173 proteins at the isoform level.

| Differential gene expression
As only six out of 54 differentially expressed transcripts on gene level between frozen and control toads were identified, the analysis herein will focus on comparisons on isoform level.We present the analysis of the gene level in the electronical supplementary material and Table S5).Isoform-level analysis revealed 277 transcripts that were differently expressed (Figure 4a-c, Figure A5 and Table S6).Using isoform-level analysis, exposure to freeze resulted in downregulation of 136 (i.e., 84 annotated) transcripts and upregulation of 141 (i.e., 89 annotated) transcripts relative to control animals (Figure 4a-c).

| Functional classification of differentially regulated genes
The biological process-oriented functional classification identified 26.9% of isoforms associated with cellular processes (Figure A5).Exposure to freeze resulted in differential expression of isoforms related to several GO slim Biological Process categories including amino-acid biosynthesis, carbohydrate metabolism and particularly gluconeogenesis as well as lipid metabolism, and stress (Figure 4a-c and Table S6).
Among the upregulated gene isoforms in toads exposed to freeze were terms associated with carbohydrate and/or lipid metabolism were cryptochrome-2 (CRY2) and insulin-like growth factor-binding protein 4 (IGFBP4).Similarly, a significant increase in the expression of aspartate aminotransferase (GOT1), involved in amino-acid biosynthesis, urea cycle, and gluconeogenesis, was shown in frozen toads compared to the control animals.Another notable upregulated isoform was adenylate cyclase type 9 (ADCY9), known to contribute to signaling cascades activated by corticotropin-releasing factor, corticosteroids and thus, involved in the neuroendocrine response to stress.

| Blood glucose level
Mean blood glucose level was 83.51% higher in frozen toads compared to the control group (U = 0.05; p = .061;N = 6; Figure 4d).

| The toll of warmer winter days: Increased metabolic demands and reduced cold tolerance
In ectothermic animals such as amphibians, metabolic rate and thus, energy demands increase with ambient temperature (Angilletta Jr et al., 2002).This may have implications for animal health and survival, particularly when resources are limited, such as during winter.
Here, we observed that toads exposed to a 4°C warmer temperature had increased metabolic rates and higher daily energy demands, leading to a faster depletion of energy budgets over time compared to toads kept at standard winter conditions (i.e., 4°C).
Across vertebrates, increased energy demands resulting from homeostatic challenges such as adverse environmental conditions are met through the release of glucocorticoid hormones (i.e., CORT in amphibians; Glennemeier & Denver, 2002) via the activation of the neuroendocrine stress axis/hypothalamic-pituitary-adrenal/ interrenal (HPA/I) axis (Crespi et al., 2013;Sapolsky et al., 2000).
Glucocorticoid hormones play a pivotal role in the metabolism of most energy reserves, regulating glucose, fat, and protein metabolism (Crespi et al., 2013;Romero & Beattie, 2022).In response to increases in metabolic rate, a main role of these hormones is to increase circulating glucose levels at a rate that corresponds to metabolic demands (Kirschman et al., 2017;Sapolsky et al., 2000).Thus, higher metabolic activity is suggested to be concomitant with increases in CORT levels (Jimeno & Verhulst, 2023).Indeed, in toads kept at a 4°C warmer temperature, we observed higher CORT release rates, suggesting that the temperature-induced higher energy demands observed therein was met through the mobilization of energy reserves mediated by an increased secretion of the glucocorticoid hormone.Furthermore, energy storages decreased withing the 48 h of exposure to higher temperatures, indicating that these reserves were mobilized to meet the increased energy demands.
Consequently, warmer winter days will result in changes in overall energy expenditure during overwintering, leading to accelerated depletion of energy reserves for all ectotherms that rely on internal energy reserves, not just amphibians (Arambourou & Stoks, 2015;Reading, 2007;Wilsterman et al., 2021).Here, for instance, we found that the fat body, as the major internal energy reserves in overwintering amphibians, would be depleted 5.26 days earlier in toads kept at higher temperatures.Despite the significant reduction in survival probabilities during overwintering and at the beginning of the breeding season upon leaving the overwintering shelter because of starvation, depleted energy reserves may further reduce immune system functioning and fecundity.This, coupled with elevated CORT levels, might increase their vulnerability to other stressors (Benard, 2015;Reading & Clarke, 1995;Rollins-Smith, 2017;Rollins-Smith & Woodhams, 2012).
In our study, cold tolerance was remarkably lower in toads kept at a 4°C warmer temperature, presumably linked to the change of energy expenditure experienced by these toads as well as the onset of physiological acclimation to higher temperatures (Lavy et al., 1997;Pörtner et al., 2000;Williams et al., 2015).Cold tolerance and energetics are closely related, as a decrease in energy reserves reduces the ability to respond effectively to cold stress, given that those energy reserves are required to cover the energy expended during cold stress exposure (Marshall et al., 2020).In freeze-tolerant species, carbohydrate energy reserves are essential for survival of freezing itself in their function as cryoprotectants (Storey & Storey, 1985, 2017).Our results are in line with findings for other ectothermic species with a life-stage specific tolerance to winter conditions.For example, Sobek-Swant et al. (2012) found that warmer winter temperatures resulted in the loss of winter acclimatization in the emerald ash borer (Agrilus planipennis), thereby increasing their vulnerability to cold spells.Furthermore, in the freeze-tolerant wood frog (Lithobates sylvaticus), repeated freezethaw due to increased winter temperature variability reduced energy reserves and thereby cold tolerance (Sinclair et al., 2013).
Therefore, changing thermal winter conditions are likely to result in a mismatch between the occurrence of extreme thermal events such as sudden freeze and the physiological mechanisms to tolerate them (Bale & Hayward, 2010;Sinclair et al., 2013;Williams et al., 2015).As a result, ectotherms including amphibians might be more susceptible to cold stress.

| Toads on ice: Limited physiological responses to sudden freeze
Amphibians inhabiting seasonal cold climates have developed a variety of winter survival strategies such as overwintering underwater or underground.However, some species that overwinter at or near the soil surface, have evolved a freeze tolerance as an adaptive cold hardiness strategy that allows surviving harsh thermal conditions (rev. in Storey & Storey, 2017).When exposed to freezing temperatures, these amphibians can synthesize and accumulate high concentrations of cryoprotectants such as glucose, glycerol, or urea in their tissues (rev. in Storey & Storey, 2017).
For example, the synthesis of glucose in L. sylvaticus is triggered by ice crystallization on the skin followed by a homeostatic stress response that stimulates glycogenolysis in the liver and its export to all other organs, resulting in a hyperglycemic state (Storey & Storey, 1985).To that end, carbohydrate energy reserves (i.e., glycogen) are mobilized and enzymes associated with liver glycogenolysis are upregulated (Storey, 1987).In our study, the blood glucose level was indeed 83% higher in frozen toads.Even if this difference was not significant, probably due to a low statistical power resulting from a small sample size, this result suggests the mobilization of carbohydrate reserves.In addition, we found evidence of transcriptional regulation of glucose biosynthesis pathways.However, contrary to our expectations, that response did not entail enhanced expression of gene isoforms directly promoting glucose synthesis via hepatic glycogenolysis (Storey, 1987).
Those enzymes, including glycogen phosphorylase (GP) and glucose-6-phosphatase (G6PC), did not show differential expression in frozen toads, and glucose transporters required for the export of glucose from the liver such as GLUT2 (Storey, 1987;Storey & Storey, 2017), were not responsive to freezing conditions in this study.Furthermore, in the pathway for glucose biosynthesis through gluconeogenesis, fructose-1,6-bisphosphatase 1 (FBP1), as one of the key enzymes, was diminished in frozen toads (Storey, 1987;Storey & Storey, 2017).Nonetheless, we found that other notable gene isoforms associated with glucose metabolism and homeostasis, mostly in response to starvation, were upregulated in frozen toads.For example, aspartate aminotransferase (GOT1) regulates the uptake of glutamine by the liver, thereby fueling the synthesis of glucose through gluconeogenesis (Just et al., 2020).Moreover, glutamine-fructose-6-phosphate aminotransferase 1 (GFPT1) was downregulated in response to freezing.Downregulation of GFPT1 might contribute to glucose availability for accumulation as it prevents the flux of glucose from the anaerobic energy production pathway to the hexosamine pathway (Denzel & Antebi, 2015).These findings suggest a response to starvation/fasting or reduced glucose availability (Just et al., 2020;Nakajima et al., 2023) to guarantee the availability of glucose for cryoprotective purposes.This might be further supported by adenylate cyclase type 9 (ADCY9), an enzyme involved in signaling cascades of the neuroendocrine stress axis and thus, contributing to the role of CORT in the regulation of circulating glucose levels to meet energetic demands (Kirschman et al., 2017;Sapolsky et al., 2000).Therefore, our results indicate that exposure to sudden freeze led to physiological responses to maintain homeostasis as well as an increase of cold hardiness by the mobilization of the cryoprotectant glucose.In contrast, we could not find any evidence for the synthesis of glycerol or urea as additional cryoprotectants as, for example, N-acetylglutamate synthase gesting that both the increases in mean temperatures as well as a period of warm winter days will make amphibians more sensitive to extreme thermal events, which are increasing in frequency as a F I G U R E A 2 Gene ontology terms describing the differential expressed genes that were up-and downregulated in frozen toads in Bufo bufo.Data shown are percentages of total genes annotated.Most genes were associated with the cellular anatomical entity (41.2%).The most common molecular functions of genes were transporter activity (9.8%) and binding (31.4%).The biological process-oriented functional classification of the annotated transcriptome identified 21.9% of genes associated with cellular processes.Furthermore, 15.6% of genes were categorized as related to stimulus responses and 19.8% to biological regulation.F I G U R E A 4 Biological process categories of genes differentially expressed in Bufo bufo compared to the control group.Data show the number of differently expressed genes that were upand downregulated in frozen toads.Genelevel analysis revealed 54 transcripts that were differently expressed between frozen and control toads.Exposure to freeze resulted in downregulation of 32 (i.e., 2 annotated) transcripts and upregulation of 20 (i.e., 3 annotated) transcripts relative to control animals.

SMI=
Individual mass × Mean SVL of population Individual SVL slope of regression logMass∼logSVL a sample concentrator (Stuart sample concentrator, SBHCONC/1; Cole-Parmer) under a fine N 2 stream at 45°C using a block heater (Stuart block heater, SBH130D/3; Cole-Parmer).Dried samples were stored at −25°C until Enzyme-Immunoassay analysis (EIA).The dried samples were re-suspended in a total volume of 500 μL (C.Gabor; personal communication) consisting of 5% ethanol (95% lab grade) and 95% EIA buffer.After re-suspension, samples were frozen at −25°C until vel measurements of hormonal levels via EIA, which took place in May 2022.
at respective acclimation temperature (i.e., 4°C and 8°C) following the calibration protocol of the manufacturer.A temperature probe enabled temperature compensation for dissolved oxygen measurements.The O 2 concentration was recorded every second and measured as O 2 % a.s.Oxygen consumption was measured for 60 min in each animal.Empty (control) chambers were run simultaneously in every trial and values were adjusted accordingly.All trials were conducted between 2000 and 0100 h to avoid circadian effects.At the end of the oxygen consumption measurements, each animal was placed back in its individual container and measurements of cold tolerance followed.Oxygen consumption in O 2 % a.s. was converted to vol % O 2 and standardized by dividing by the body mass of each individual, resulting in SMR units being mL O 2 × h −1 × g −1 .The analyses were performed in PreSens Oxygen Calculator Software (PreSens Precision Sensing GmbH).In order to determine differences in the energy expenditure of the animals under the different acclimation temperatures, DEE in J × (d × g) −1 was calculated from the standardized mean of the metabolic rates per hour and gram of body mass using the caloric equivalent and assuming a respiratory quotient of 0.85 (resulting in 20.37 J × mL O 2 −1 ).Finally, we calculated the maximum survival time in d if toads actually relied on the energy storages of the fat bodies to cover their DEE (i.e., energetic maintenance costs) (see Section 2.3.3).

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I G U R E 2 (a) Mass (g) and (b) SMI as a measure of body condition of adult B. bufo before and after a 48-h acclimation period at 4°C and 8°C.Boxes and whiskers show 25th to 75th and 10th to 90th percentiles, respectively; black lines indicate the median.Dot = single data points.Blue boxes = toads acclimated to 4°C.Orange boxes = toads acclimated to 8°C.Gardner-Altman estimation plots displaying the mean difference between acclimation temperatures on (a) log-transformed cold tolerance (CT min ,°C) in adult B. bufo (N = 30).(b) Start of CT min trial.(c) Toad at CT min .Boxes and whiskers show 25th to 75th and 10th to 90th percentiles, respectively; black lines indicate the median.Dot = single data points.Blue box = freeze group.Orange box = control group.Furthermore, 13.8% of isoforms were categorized as related to metabolic processes and 14.2% to biological regulation.The most common molecular functions of isoforms were catalytic (26.2%) and binding (27.4%).Most of isoforms were associated with the cellular anatomical entity (53.9%).

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I G U R E 4 (a) Heatmap with hierarchical clustering of genes (rows) showing relative expression of all significant differentially expressed gene isoforms (DEGs) (p < .01) in individual control group (N = 5, left column) and individual freeze group (N = 5, right column) livers.Orange indicates upregulation, blue indicates downregulation.(b) Volcano plot showing DEGs in transcriptomes of control group and freeze group livers.The logFC in counts for each transcript (magnitude of differential expression) are plotted on the x-axis against the statistically significant −log10 (p adj < .01)values on the y-axis.Dashed lines correspond to p adj and logFC cut-off values set at p < .01 and 2.0, respectively.Gray dots indicate DEGs (p adj < .01);orange and blue dots indicate DEGs (p adj < .01)above or below the set logFC threshold of +2.0 or −2.0, respectively.(c) Biological process categories of isoforms differentially expressed in Bufo bufo compared to the control group.Data show the number of/percentage of differently expressed gene isoforms that were up-and downregulated in frozen toads.(d) Effect of freeze exposure on blood sugar level (mg × dL −1 ).Boxes and whiskers show 25th to 75th and 10th to 90th percentiles, respectively; black lines indicate the median.Dot = single data points.Blue box = freeze group (n = 3).Orange box = control group (n = 3).

(
NAGS), known to be involved in the urea cycle, was downregulated.Measurements of urea could be included by future studies(de Amaral et al., 2023).Consequently, B. bufo might be capable of at least a short-term freeze tolerance which might provide a means in fitness if toads encounter a sudden night freeze, for instance, on their way to their breeding sites after leaving the overwintering shelter.Nonetheless, the mobilization of glucose for cryoprotective purposes requires energy reserves (i.e., carbohydrates for glycogenolysis and proteins for gluconeogenesis).If these energy reserves are already drained due to the catabolic effect of increased mean winter temperatures with climate change(Marshall et al., 2020), cryoprotective accumulation of glucose might be constrained with consequences for animal survival in false springs events.Future research is needed to better understand the interplay of elevated mean temperatures and an increased frequency of extreme cold events on amphibian physiological resilience.This research is crucial to unraveling the potential double jeopardy of winter climate change, which may have been underestimated in its role in the decline of temperate amphibian populations.5 | CON CLUS IONEnergy is the only currency that counts in and for life, and climate change has the potential to disrupt energy balances throughout the annual cycle in several ways such as through increased energetic demands and fewer available resources.Consequently, wildlife might become more susceptible to environmental stress, particularly when coping mechanisms are limited.Our study points for the first time to the complex physiological challenges that amphibians face as a result of changing thermal conditions due to winter climate change.Overall, our results highlight that warmer winters and repeated freeze-thaw cycles impose physiological costs that could reduce energy reserves and might thereby affect amphibian health and survival.Moreover, we demonstrated that warmer winter temperatures reduced cold tolerance, sug-

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I G U R E A 3 (a) Heatmap with hierarchical clustering of genes (rows) showing relative expression of all significant differentially expressed genes (DEGs) (p < .01) in individual control group (N = 5, left column) and individual freeze group (N = 5, right column) livers.Orange indicates upregulation, blue indicates downregulation.(b) Volcano plot showing DEGs in transcriptomes of control group and freeze group livers.The logFC in counts for each transcript (magnitude of differential expression) are plotted on the x-axis against the statistically significant −log10 (p adj < .01)values on the y-axis.Dashed lines correspond to p adj and logFC cut-off values set at p < .01 and 2.0, respectively.Gray dots indicate DEGs (p adj < .01);orange and blue dots indicate DEGs (p adj < .05)above or below the set logFC threshold of +2.0 or −2.0, respectively.Gene-level analysis revealed 54 transcripts that were differently expressed between frozen and control toads.Exposure to freeze resulted in downregulation of 32 (i.e., two annotated) transcripts and upregulation of 20 (i.e., three annotated) transcripts relative to control animals.
Non-parametric Mann-Whitney U-tests were used to compare CORT release rates, SMR, DEE, and CT min between toads acclimated to 4°C and 8°C.Paired-samples t-tests were employed to assess the difference in body mass and SMI at collection and at the time of measurements (i.e., after 48 h) of each individual.
After the 48-h acclimation period, half of the toads (n = 5; freeze group) were transferred to a portable freezer (DOMETIC Coolfreeze CDF 36, Dometic Germany GmbH) and kept at a mean temperature of −2°C for 6 hours to simulate a night freeze.The other half of the toads remained at 4°C (n = 5; control group).After 1, 3, and 6 h, we quickly measured T cloaca of all animals by inserting a temperature probe into the animal's cloaca in their respective acclimation environment (Figure